MasterProof

The Proton-Driven Rotor of ATP Synthase: Ohmic Conductance (10 fS),and Absence of Voltage Gating

Boris A. Feniouk,*y Maria A. Kozlova,* Dmitry A. Knorre,*y Dmitry A. Cherepanov,*z

Armen Y. Mulkidjanian,* and Wolfgang Junge**Division of Biophysics, Faculty of Biology/Chemistry, University of Osnabruck, Osnabruck, Germany; yA.N. Belozersky Institute ofPhysico-Chemical Biology, Moscow State University, Moscow, Russia; and zInstitute of Electrochemistry,Russian Academy of Sciences, Moscow, Russia½AQ1�

ABSTRACT The membrane portion of F0F1-ATP synthase, F0, translocates protons by a rotary mechanism. Protonconduction by F0 was studied in chromatophores of the photosynthetic bacterium Rhodobacter capsulatus. The discharge ofa light-induced voltage jump was monitored by electrochromic absorption transients to yield the unitary conductance of F0. Thecurrent-voltage relationship of F0 was linear from 7 to 70 mV. The current was extremely proton-specific (.107) and varied onlyslightly (�threefold) from pH 6 to 10. The maximum conductance was �10 fS at pH 8, equivalent to 6240 H1 s�1 at 100-mVdriving force, which is an order-of-magnitude greater than that of F0F1. There was no voltage-gating of F0 even at low voltage,and proton translocation could be driven by DpH alone, without voltage. The reported voltage gating in F0F1 is thus attributableto the interaction of F0 with F1 but not to F0 proper. We simulated proton conduction by a minimal rotary model including therotating c-ring and two relay groups mediating proton exchange between the ring and the respective membrane surface. Thedata fit attributed pK values of �6 and �10 to these relays, and placed them close to the membrane/electrolyte interface.

INTRODUCTION

ATP synthase or F0F1-ATPase, a key enzyme of bio-

energetics, is present in the photosynthetic and respiratory

coupling membranes of eubacteria, mitochondria, and chlo-

roplasts. The enzyme is bipartite. Its membrane-embedded

F0-portion transports protons down the electrochemical

gradient (DuuuH1) and generates torque. The peripheral F1-

portion synthesizes ATP at the expense of this mechanical

energy. Both F0 and F1 are rotary motors/generators and

steppers. They are mechanically coupled by a central rotary

shaft and held together by a peripheral stator (for general

reviews, see Senior et al., 2002; Noji and Yoshida, 2001;

Leslie andWalker, 2000; Boyer, 1997; Junge et al., 1997; and

for the sodium ATP synthase, see Dimroth, 2000; Dimroth

et al., 1999). By its rotary mechanism F0 is distinguished from

most other proton or cation transporters with the exception of

the structurally unrelated ionic drive of the bacterial flagellar

motor.

F0 is composed of three types of subunits in a stoichiom-

etry of a1b2c10–14. All three types of subunits are required

for proton conduction (Schneider and Altendorf, 1987;

Altendorf et al., 2000). Multiple copies of the hairpin-shaped

c-subunit form a ring in the membrane to which subunits

a and b are attached from the outer side (Lightowlers et al.,

1988; Angevine and Fillingame, 2003; Fillingame and

Dmitriev, 2002; Jiang et al., 2001; Muller et al., 2001;

Seelert et al., 2000; Stock et al., 1999). ATP hydrolysis in

F0F1 is coupled to the rotation of the c-ring (Panke et al.,

2000; Sambongi et al., 1999; Tsunoda et al., 2000, 2001),

and power is elastically and without slip transmitted between

the drive in F1 and the ring of F0 (Cherepanov et al., 1999;

Cherepanov and Junge, 2001; Panke et al., 2001).

The currently favored mechanism of torque generation by

proton flow (Junge et al., 1997; Elston et al., 1998) has been

based on the following assumptions: Brownian rotary

motion of the c-ring relative to subunit a, two non-co-linear

access channels for protons from either side, and two

electrostatic constraints, namely the acid Glu residue (Asp in

Escherichia coli) in the middle of the hairpin of c being

deprotonated when in contact with subunit a and protonated

when facing the core of the membrane (Junge et al., 1997;

Elston et al., 1998). Modifications to this basic scheme have

been formulated to account for Na1-translocation in F0 from

Propionigenium modestum (Dimroth et al., 1999) or to

account for internal librational motion in the ring (Fillingame

et al., 2000). The basic principle of this rotary mechanism of

proton conduction—Brownian fluctuations and two electro-

static constraints—has remained (see Junge, 1999). It is

obvious that a comprehensive understanding of rotary proton

conduction requires both structural and kinetic information.

Most recent studies on this enzyme have focused on the

F1-portion, whereas most studies on the mechanism of

proton conduction by F0 date back by more than one decade.

The assessment of the conductance of F0 has been hampered

by complications:

1. The conductance is too small for the single-molecule

patch-clamp approach; see below for one report to the

contrary.

Submitted December 9, 2003, and accepted for publication February 11,2004.

Address reprint requests to Wolfgang Junge, Abt. Biophysik, FB Biologie/

Chemie, Universitat Osnabruck, D-49069, Osnabruck, Germany. Tel.:

49-541-969-2872; Fax: 49-541-969-2262; E-mail: junge@uos.de.

� 2004 by the Biophysical Society

0006-3495/04/06/1/16 $2.00 doi: 10.1529/biophysj.103.036962

Biophysical Journal Volume 86 June 2004 1–16 1

MasterProof

2. Studies with large ensembles suffer from the difficulty

in determining the proportion of F0 that is actually

conducting.

3. Studies with reconstituted F0 using glass electrodes for

recording pH transients have often lacked sufficient time

resolution.

Thus it is not surprising that the reported values for the

conductance at neutral pH diverged by four orders of

magnitude. Some studies on the conductance of F0 have

yielded rather low values in the range of 0.1 fS, which is

equivalent to a rate of 62 s�1 at a driving force of 100 mV

(Negrin et al., 1980; Friedl and Schairer, 1981; Schneider

and Altendorf, 1982; Sone et al., 1981; Cao et al., 2001).

This value would be insufficient to cope with the turnover

rate of the coupled F0F1 enzyme. For chloroplast CF0 we

have found a 100-fold-higher figure, 9 fS (or 5600 s�1 at 100

mV) under the assumption that all exposed F0 molecules

contributed to the relaxation of the transmembrane voltage

(Schonknecht et al., 1986). This conductance is compatible

with the maximum proton turnover rate of the coupled

enzyme, �1000 s�1. Later we obtained circumstantial evi-

dence for a majority of inactivated F0, which raised the

conductance to a value of 1 pS (Lill et al., 1986; Althoff et al.,

1989). Such high rates by far exceeded the diffusive proton

supply to F0, as then noted, and it has remained questionable

whether the underlying assumption was correct. This work

has also provided evidence for an extreme proton specificity

of the conductance (.107 over other cations), small pH-

dependence between 5.5 and 8, a H/D-kinetic isotope effect

of 1.7, and an activation energy of 42 kJ/mol in H2O and 47

in D2O. The latter data were not affected by the above

ambiguity over the absolute magnitude of the conductance

(Althoff et al., 1989).

A conductance of similarly highmagnitude was reported for

CF0CF1 in a lipid bilayer formed by the dip-stick technique,which is related to patch-clamp (Wagner et al., 1989). Gated

single channel currentswere observed. They peaked at 0.55 pA

at 180 mV, implying a conductance of 0.4 pS, and channels

were gated open with a sharp activation above 100 mV. The

authors attributed this voltage-gated conductance to the proton

(Wagner et al., 1989). Whether this attribution was correct has

remained an open question, in particular because proteolipo-

somes, which contained the purified c-subunit alone, haverevealed unspecific cation channels (Schonknecht et al., 1989;

for the proton conduction at pH 2 of the purified c-subunit seeSchindler and Nelson, 1982).

The lack of information on the number of active F0‘‘motors’’ per membrane area was a major obstacle for

obtaining a reliable estimate of its proton conductance. Our

recent studies on isolated chromatophore vesicles from the

photosynthetic bacterium Rhodobacter capsulatus (Feniouk

et al., 2001, 2002) havepaved away toovercome the ambiguity

over the proportion of active and inactive F0. The key was to

prepare vesicles of such small size as to contain less than one

copy of F0 on the average. Then, any rapid relaxation of the

transmembrane voltage could be attributed to the subset of

vesicles containing mainly a single F0 molecule.

In this work we prepared vesicles with 28-nm mean

diameter as checked by transmission electron microscopy

(TEM). Excitation of a suspension of such vesicles with

a flash of light generated a voltage step across the membrane.

The magnitude of the initial voltage jump after a saturating

flash of light (70 mV) was calibrated by comparison with

electrochromic absorption transients that were induced by

submitting vesicles from the same batch to a salt-jump. The

relaxation of the voltage was monitored by electrochromic

absorption transients of intrinsic carotenoids, and the proton

flow by pH-indicating dyes. The relaxation was biphasic.

The fast phase (25%) was mediated by F0 as evident from its

sensitivity to the F0-inhibitor oligomycin. It was mainly

attributable to proton conduction in the subset of vesicles

containing a single copy of F0.

MATERIALS AND METHODS

Cell growth and preparation of stockchromatophores

Cells of Rb. capsulatus (wild-type, strain B10) were grown photohetero-

trophically on malate as the carbon source at 130�C (Lascelles, 1959) as

described elsewhere (Feniouk et al., 2001). The cells were disrupted by

sonication with a W-250 Branson Sonifier (Branson, Soest, The Nether-

lands); the relative power output was set at 35%. The material was exposed

to one series of five exposures for 20 s with 1-min incubation on ice in be-

tween. Large cell fragments were removed by centrifugation (20,000 3

g, 20 min, 4�C). The supernatant was collected and centrifuged again

(180,000 3 g, 90 min, 14�C). The pellet was resuspended in 20 mM

glycylglycine-KOH (pH 7.4), 5 mM MgCl2, 50 mM KCl, 10% sucrose. It

contained chromatophores at a bacteriochlorophyll concentration of 0.3–

1.0 mM. These stock chromatophores were stored at �80�C until use. The

concentration of bacteriochlorophyll in the samples was determined

spectrophotometrically in acetone-methanol extract at 772 nm according

to Clayton (1963). The amount of functionally active photosynthetic

reaction centers (RCs) was estimated from the extent of flash-induced

absorption changes at 603 nm (Mulkidjanian et al., 1991). The above

protocol was used throughout the experiments, except for the data shown in

Fig. 4 B and Fig. 5, where chromatophores from another stock prepared with

a weaker sonifier were used. However, the F1 depletion was performed in the

same way as for the other batches. The smallness of the F1-depleted vesicles

(see below) resulted from the sonication during the F1 depletion.

Preparation of F1-depleted chromatophores

Chromatophores from the stock were thawed, diluted 25-fold with 1 mM

EDTA, and titrated with KOH to pH 8, in daylight (Melandri et al., 1970,

1971; Baccarini-Melandri et al., 1970). The suspension was submitted to

another series of sonications (W-250 Branson, relative output 35%), five

times for 20 s each with 1-min pausing on ice, and centrifuged at 180,000 3

g, 90 min, 14�C. The pellet was resuspended in a medium containing 20

mM glycylglycine-KOH (pH 7.4), 5 mM MgCl2, 50 mM KCl to yield

a bacteriochlorophyll concentration of 0.3–1.0 mM and stored at �80�Cuntil use. For measurements of the pH transients F1-depleted chromato-

phores were prepared without glycylglycine-KOH.

2 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

Chromatophore size determination by TEM

Chromatophores were suspended in a medium with 100 mM KCl, 20 mM

glycylglycine, 5 mM MgCl2, pH 8.0 to yield a bacteriochlorophyll

concentration of 5 mM. A drop of the suspension was applied on the

carbon-coated grid, stained with 3% uranyl acetate, washed with water, dried

and examined in a Zeiss 902 transmission electron microscope (Carl Zeiss,

Oberkochen, Germany). F1-depleted chromatophores appeared as disks with

diameters of 39.7 6 1.0 nm (SD 7.9 nm; see Results; also see Discussion).

Flash spectrophotometry

The kinetic flash-spectrophotometer was described elsewhere (Feniouk et al.,

2001). The bandwidth of the measuring beam, 8 nm, was set by interference

optical filters (Schott, Mainz, Germany). Changes in transmitted light

intensity (DI) were monitored by a photomultiplier (Thorn EMI, 9801B,

UK) shielded from the actinic flash with two blue filters (BG 39, 4 mm,

Schott). The electrical bandwidth was 3 kHz and the digital time-per-address

of the averager was 200 ms. The optical path was 1 cm, both for the exciting

and for the measuring beam. The final concentration of bacteriochlorophyll

in the cuvette was 8–12 mM.

Transient signals were generated by flashing light at a repetition rate of

0.08 Hz. Eight signals at 522 nm were averaged in recordings of the

electrochromism and 32 inmeasurements with pH indicators. During the dark

time (12.5 s) between flashes the electrochromic signal decayed to,5%of its

initial value after the flash of light, even if the conductance of F0 was blocked

by oligomycin (not documented). The actinic flashes were provided by

a xenon flash lamp (full-width at half-maximum ¼ 10 ms) with a red optical

filter (RG 780, Schott); the total energy density on the cuvette (not attenuated)

was 12mJ cm2. The peaks of the xenon emission spectrum (e.g., 825 and 875

nm) overlap with regions of low absorption of the chromatophores (their

peaks are at 800 and 850 nm). We calculated an effective optical density for

the excitation by convoluting the xenon emission and chromatophore

absorption. With the highest concentration (12 mM bacteriochlorophyll) in

the cuvette it was 0.6 OD, one-half of the value (1.2 OD), which was

calculated for the peak absorption at 850 nm. In control experiments with

lower bacteriochlorophyll concentration (6mM), the effective optical density

was 0.3, which provided a rather homogenous excitation profile over the

thickness of the optical cell. Neutral density filters were used to attenuate the

actinic light flash when indicated (see Fig. 5 legend).

Electrochromic carotenoid bandshifts at 522 nmwere used as a molecular

voltmeter to monitor Dc (Junge and Witt, 1968; Clark and Jackson, 1981;

Symons et al., 1977; Jackson and Crofts, 1971).

Transients of the pH inside chromatophores (DpHin) were monitored by

absorption changes of the amphiphilic pH indicator Neutral red at 545 nm as

in Auslander and Junge (1975) and Mulkidjanian and Junge (1994). In these

experiments 0.3% BSA was used as an impermeable buffer to quench pH

changes in the suspending medium (Auslander and Junge, 1975). pH

transients in the outer medium (DpHout) were monitored by absorption

changes of the hydrophilic pH indicator Cresol red at 575 nm (Saphon et al.,

1975) and calibrated in pH units. For each pH indicator, control traces

without the indicator were recorded and subtracted.

Calibration of the carotenoid bandshift inmillivolts of transmembrane electricalpotential difference

Chromatophores were suspended in a medium containing 20 mM

glycylglycine (pH 8.0), 5 mM MgCl2, 2 mM NaCN, 3 mM myxothiazol,

3 mM oligomycin, and KCl. The KCl concentration was varied, and NaCl

was added to keep the total concentration of KCl1NaCl constant at 250

mM. Valinomycin (1 mM stock solution in EtOH, final concentration in the

cuvette 1.6 mM) was added to induce a diffusion potential of K1; the

resulting absorption transients at 522 nm were recorded in the same

instrument used for flash spectrophotometry.

RESULTS

Vesicle diameter and transmembrane voltageof F1-depleted chromatophores

Fig. 1 A shows the size-distribution of F1-depleted chrom-

atophores as obtained by TEM. The diameter of the circular

spots, 39.7 6 1.0 nm (SD 7.9 nm), was attributable to

vesicles that were flattened by drying on the microscope grid.

If these were swollen to spherical shape the expected

diameter was 28.1 6 0.7 nm (SD 5.6 nm).

Fig. 1 B shows the extent of the absorption transients at

a wavelength of 522 nm, the peak of the electrochromic

transient, as a function of the KCl concentration in the

suspending medium. These transients represent the elec-

trochromic response of intrinsic pigments to the diffu-

sion potential of K1, which was generated by suspending

FIGURE 1 (A) Size distribution of F1-depleted chromatophores by

transmission electron microscopy (TEM). Because of drying on the

microscope grid the originally spherical particles appeared as flat disks

(see text). (B) Absorption transients attributable to carotenoid bandshifts of

electrochromic origin as function of the K1 concentration in the suspending

medium. These transients were caused by a diffusion potential that was

generated by submitting chromatophores to a K1-concentration jump in the

presence of the K1-ionophore valinomycin. F1-depleted chromatophores

(940 mM bacteriochlorophyll) were preincubated with 45 mMKCl, 205 mM

NaCl, 5 mM MgCl2, 20 mM glycylglycine-NaOH, pH 7.9. The final

bacteriochlorophyll concentration in the samples was 12 mM.

Proton Conductivity of F0 3

Biophysical Journal 86(6) 1–16

MasterProof

F1-depleted and KCl-loaded chromatophores in a KCl-

containing buffer and adding the potassium-specific iono-

phore valinomycin. The photometric response was linearly

related to the logarithm of the KCl concentration over two

decades. In the experiment documented in Fig. 1 B there was

no electrochromic transient induced at a K1 concentration of

35 mM. Apparently, 35 mM was the effective concentration

of KCl in the chromatophore lumen in this particular

experiment. The linear dependence on the ambient KCl

concentration was perfectly reproducible, whereas the

crossover point and the slope varied slightly between

experiments. We took the variation of the intensity jump

over one decade of KCl as representing 58 mV, as usual.

If the transmembrane voltage jump was generated by the

photosynthetic reaction centers (RCs) in response to a sin-

gle saturating flash of light, the electrochromic absorption

transient was composed of two contributions, one being the

response to the delocalized and transmembrane field and the

other to the local electric field originating from the oxidized

primary electron donor. In the calibration procedure, only the

first component had to be considered. We eliminated the

delocalized transmembrane component by adding valino-

mycin, 1 mM, and estimated its magnitude (see Fig. 2). The

residual valinomycin-insensitive absorption changes (�5%

of the total flash-induced jump in our sample, see Fig. 2)

relaxed in the range of 100 ms, after the reduction of primary

electron donor in the RC. Although the relative extent of this

component was only 5% in the samples studied in this work,

it can reach 15% if no redox mediator is present in the

medium.

With the above correction, the calibration yielded 706 15

mV for the RC-generated transmembrane voltage jump after

a saturating flash of light (seven calibration experiments on

three different preparations of F1-depleted chromatophores).

We estimated the number of RCs in these vesicles. The

estimate was based on the voltage of 70 mV, on the usual

figure for the specific capacitance of biomembranes, C ¼1 mF cm�2 (Hille, 1984), and on the mean vesicle radius of

14 nm, as determined by TEM. The capacitor equation reads:

n 3 eo ¼ A 3 C 3 DU, wherein n denotes the number of

RCs per vesicle, eo, the elementary charge, A the vesicle area,

and DU the transmembrane voltage. We obtained a value of

n ¼ 11 as the mean number of RCs in our preparation of

F1-depleted vesicles. This number implied a surface density

of½AQ2� 7.3 � 10�9 mol m�2, �40% larger than the previously

determined 5.3 � 10�9 mol m�2 (Packham et al., 1978).

Electrochromic absorption transients with andwithout active cytochrome bc1 complex

Fig. 2 A shows flash-induced absorption transients at 522 nm

in F1-depleted chromatophores. The rapidly decaying lower

trace was obtained with valinomycin. It reveals the

magnitude of the electrochromic response to the trans-

FIGURE 2 Electrochromic absorption transients at a wavelength of 522

nm in F1-depleted chromatophores as caused by a short flash of light. The

upper traces were obtained without and with blocking of F0 by 3 mM

oligomycin, respectively. The difference trace (6 oligomycin) represents thecharge flow across F0 (A) without and (B) with 3 mM myxothiazol added to

deactivate the electrogenic and proton pumping activity of the cytochrome

bc1 complex. The actinic flash is indicated by an arrow. The bottom traces

were obtained in the presence of 1 mM valinomycin to enhance the ion

conductance of all chromatophores in the sample. It was apparent that

absorption transients not originating from electrochromism as well as any

electrochromic responses to localized electric fields were negligible under

our conditions. The suspending medium contained: 100 mM KCl, 5 mM

magnesium acetate, 2 mM K4[Fe(CN)6], 5 mM 1,1#-dimethylferrocene

(DMF), 2 mM KCN, 20 mM glycylglycine-KOH, pH 7.9; chromatophores

were added to 12 mM bacteriochlorophyll.

4 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

membrane and delocalized electric field. The upper traces

were obtained with and without added oligomycin (Linnett

and Beechey, 1979) as an inhibitor of proton conduction

through F0. The slowly rising lower trace gives the differ-

ence between the upper two, corresponding to the cumula-

tive charge flow through F0. The absorption change after

the actinic flash was biphasic (see Fig. 2 A, trace witholigomycin added). The sharp (here not time-resolved)

increase is attributable to the charge separation in the RC.

Its magnitude is fairly independent of pH, redox potential,

and temperature (Jackson, 1988). We used the extent of the

fast rise to normalize transients that were obtained in

different samples. In Fig. 2 A the fast onset was followed by

a slowly rising phase. It was attributable to the electrogenic

reaction in the cytochrome bc1 complex (bc1; Crofts and

Wraight, 1983; Jackson, 1988). This slow phase was

abolished upon the addition of the bc1-inhibitor myxothiazol

(see Fig. 2 B).The decay of the electrochromic transient was rapid if F0

was conducting and slow if it was blocked. The latter proved

the low ion permeability of the membrane. A clear-cut

interpretation of the difference trace in Fig. 2 A was

hampered by the activity of the cytochrome bc1 complex

for two reasons: 1), bc1 translocates electrons and protons,

and thereby generates a transmembrane pH jump in addition

to a voltage jump; and 2), the turnover of bc1 is hindered by

a large DuuuH1 . In other words, the turnover of bc1 is not

independent of the presence of the F0-inhibitor oligomycin.

These effects were eliminated by inactivating the bc1-complex with myxothiazol, which abolished the slow rise

of the electrochromic transient as documented in Fig. 2 B.The RCs generated a voltage step of the same magnitude as

when bc1 was active (compare Fig. 2, A and B; also see

Mulkidjanian and Junge, 1994). Only in this case does the

difference trace (6 oligomycin) reflect the relaxation of the

transmembrane voltage step in those vesicles that have

a single or, in a minor fraction only, several F0 complexes.

Comparison of the relaxation times in Fig. 2, A and B,revealed the systematic error when assessing the voltage

relaxation with active bc1 (the apparent relaxation time was

7–11 ms as compared with 2–4 ms in the presence of

myxothiazol). Put loosely, F0 first transferred the protons

driven by the RC-generated Dc and then ‘‘waited’’ for the

protons that were ejected by the cytochrome bc1-complex.

The rate of proton transfer through F0 in the presence of

myxothiazol was almost independent of the presence of

a penetrating pH-buffer glycylglycine (see Table 1).

To avoid complications coupled with slow proton

release from bc1, all subsequent studies were carried out in

the presence of the bc1-blocker myxothiazol (conditions as in

Fig. 2 B). The relative extent of rapid proton transfer

represented the fraction of vesicles containing at least one

copy of active F0. In the preparations used in this work this

fractionwas 26%.According to Poisson’s distribution,�22%

of total contained a single copy of F0, and �3% two copies.

The discharge time of the vesicles’ electric capacitance by

F0 was then apparent as the relaxation time of the difference

trace (6 oligomycin) as in Fig. 2 B. The alternative inter-

pretation of the biphasic decay of the voltage, the discharge

of the membrane’s electric capacitance by a voltage-gated

channel, was ruled out by the observation of the same

biphasic behavior after a lower starting voltage (nonsaturat-

ing energy of the exciting flash; see Fig. 3 in Feniouk et al.,

2002).

The pH-dependence, H/D-kinetic isotope effect,and Arrhenius activation energy of protonconduction by F0

Fig. 3 shows the variation of the rate constant of the electric

relaxation via F0 (in the presence of myxothiazol) as function

of the pH and the pD. ( ½AQ3�The pD was determined as pH-meter

reading plus 0.4; Glasoe and Long, 1960.) Magnesium

acetate was omitted from the measuring medium because of

precipitation at pH values .8.5. However, up to pH 8.5 the

presence of magnesium cations had no detectable effect on

the pH-dependence (data not shown). The highest rate

TABLE 1 Apparent relaxation time constants of

transmembrane proton flow through F0 at pH 8

(average values from four different trials)

Experimental conditions Time constant, ms

Control (no additions) 8.6 6 1.4

3 mM myxothiazol 3.8 6 1.1

20 mM glycylglycine, Na2HPO4, MES, glycine

each, and 3 mM myxothiazol

3.1 6 0.9

The medium contained 100 mM KCl, 2 mM KCN, 5 mM 1,1#-dimethylferrocene (DMF), 2 mM K4[Fe(CN)6].

FIGURE 3 The rate constant of charge flow through F0 (see Fig. 2 B) asa function of the pH and the isotope composition of the medium (H2O: s,

D2O: d). Medium contained 20 mM glycylglycine, 20 mM sodium

phosphate, 0 mM 4-morpholinepropanesulfonic acid (MES), 20 mM

glycine, 50 mM KCl, 2 mM K4[Fe(CN)6], 5 mM DMF, 2 mM KCN, 3

mM myxothiazol. The solid line was calculated by the kinetic model of F0conductance as described in the text with the parameters listed in Table 2.

Proton Conductivity of F0 5

Biophysical Journal 86(6) 1–16

MasterProof

constant, 350 s�1, was observed;pH 8. The small variation

of the rate in the wide pH range from 6 to 10 was astounding.

The rate constant was decreased only threefold at both the

acid and alkaline end of the pH range. The H2O/D2O-kinetic

isotope effect was as large as two at acid pH and decreased to

�1 above pH 8 (Fig. 4 A). The extent of the charge flow via

F0 was pH-independent within the experimental error (when

care was taken to avoid the aging of the preparation). The

Arrhenius activation energy of the electric relaxation was 57

6 4 kJ mol�1 at pH 8, and increased to 72 6 3 kJ mol�1

upon acidification (Fig. 4 B).

It was noteworthy that the changes of the conductance by

60 min exposure to acid (e.g., pH 5.5) and alkaline pH (pH

10.5) were reversible. The electric relaxation time constant

changed by ,10% after back-titration into the neutral pH

range.

The Ohmic behavior of F0

The following data on the electric conductance of F0 were

obtained in the presence of myxothiazol, to avoid the com-

plications caused by the electrogenic and proton pumping

activity of bc1. We changed the initial voltage jump by

varying the flash energy. With a mean number of RCs per

vesicle of 11 (see above) the initial extent of the voltage

could be varied over one order of magnitude.

Fig. 5 shows two difference transients (6 oligomycin)

reflecting the charge transfer through F0. These transients

were obtained at 100% and 33% saturation (judging from the

initial extent of the absorption jump at 522 nm), respectively.

The extent of the initial step of absorption was normalized. It

was obvious that the two transients perfectly superimposed.

The transient at nonsaturating flash energy was only dis-

cernible by greater noise from the one obtained at saturating

flash energy (circles). The relaxation was almost perfectly

monoexponential (fit curve not shown in Fig. 5) and inde-

pendent of the initial voltage jump. The same relaxation time

was found when the flash intensity was attenuated to yield

10% saturation (not documented).

The original traces in Fig. 5 were obtained at a bacterio-

chlorophyll concentration of 12 mM where the effective

optical density of the sample in the excitation region was 0.6

(see Materials and Methods). It caused a drop of the flash

energy from the entry to the exit of the cuvette (1-cm

thickness) from 100% to 25%. This did not matter for the

traces with 100% signal saturation, because the flash energy

was far-oversaturating, but it did matter for the traces at 33%.

In this respect, it was noteworthy that we obtained the same

relaxation times both in samples with halved bacteriochlo-

rophyll concentration (6 mM, optical density 0.3, flash

energy drop across the cuvette from 100% to 50%) and in

samples with 10% saturation. It justified the neglect of the

saturation profile in the following interpretation of our data.

The traces at the bottom of Fig. 5 show the difference

between the normalized 100% and 33% transients; the devia-

tion between them was very small indeed. Such a voltage-

independent relaxation characterizes the discharge of a ca-

pacitor (area-specific capacitance ¼ C/AsV�1 m�2) by an

Ohmic resistor (area-specific conductance¼ G/AV�1m�2). It

yields a voltage-independent relaxation time, t ¼ C/G. F0behaved as an Ohmic resistor, and it showed no voltage-

gating.

The rapid relaxation of the voltage (relaxation time ¼�2.9 ms at pH 8; see Fig. 3) was only observed in the small

subset of vesicles containing F0 (26% of total). The electric

capacitance of vesicles C was calculated by their size using

FIGURE 4 Effect of pH on the H/D-isotope effect and on the Arrhenius

activation energy of the charge transfer through F0. (A) H/D isotope effect

was calculated as the ratio of the rate constants in H2O and D2O (data taken

from Fig. 3). (B) The pH-dependence of the Arrhenius activation energy

(Ea). Measurements were done in the presence of 3 mM myxothiazol in the

temperature interval from 3�C to 40�C. A single experiment was done at pH

9; other points are the average of at least three experiments.

6 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

the usual figure for biological membranes, 1 mF cm�2 (Hille,

1984). The conductance of F0, g, was calculated according tog¼ A 3 G with G¼ C/t. Herein A denotes the vesicle area,

t the electric relaxation time, and G and C are the area-

specific conductance and electric capacitance of the mem-

brane, respectively.

The shortest relaxation time observed, 2.3 ms, gave rise to

a calculated conductance of 10.5 fS. It implied the transport

of 6500 protons/s at 100-mV steady driving force or of

13,000/s at 200 mV.

It was noteworthy that the proton-transporting properties

of the Rb. capsulatus F0 were robust against the changes ofthe concentration of K1, Na1, Cl�, of various permeating

and nonpermeating buffers, and redox mediators (not

documented).

The proton specificity of F0

We complemented measurements of the electric relaxation

with measurements of the pH transients in both phases—the

suspending medium (hydrophilic indicator Cresol red,

wavelength 575 nm) and the interior of chromatophores

(amphiphilic indicator Neutral red, 545 nm, with BSA as

a nonpermeating buffer). The results are documented in Fig.

6. The data were obtained with active bc1, to demonstrate the

proton uptake from the bulk medium by the RC and by bc1(Fig. 5, A and B) and the proton release into the interior of

chromatophores by bc1 (Fig. 5, C and D). The alkalization ofthe bulk and the acidification of the interior were fully ap-

parent when F0 was blocked by oligomycin (traces 2), andboth were diminished when F0 was conducting (traces 1). Therespective difference traces gave the F0-related pH-relaxation,

however, with somewhat distorted kinetics because of the

slow proton-pumping activity of bc1 (see above discussion inthe context of Fig. 2, A and B). Ignoring the complex kinetics

of these difference transients, they proved that the major ion

transported by F0 was the proton at an ambient pH 7.9 against

a background of 100 mM KCl. Similar results were observed

with NaCl and LiCl (data not shown). In other words, the

proton selectivity of F0 was at least 107.

FIGURE 5 Transient charge transfer through F0 in the

presence of 3 mMmyxothiazol as inhibitor of cytochrome-

bc1. (A) Two transients of charge flow through F0, both

electrochromic differences (6 oligomycin) as the bottom

trace in Fig. 2 B.s, the transient recorded under excitation

with a flash of saturating energy. Noisy line, the transient

recorded at attenuated excitation flash energy to produce

33% signal saturation. The extent after the flash was

normalized to unity. The insert maps allowed (uncolored)

and forbidden (hatched) area in the parameter-field of a

and b (see details in the text). (B) The noisy line represents

the difference between the original transients shown in A;a histogram of the deviations is shown in the insert. The

thick solid line is the same difference calculated according

to the model with the parameters a ¼ 0.05, b ¼ 0.71, g ¼10 (other parameters are listed in Table 2; see details in the

text). It is consistent with the experimental error of 3%.

The dashed curve was calculated with the same set of

parameters except that g ¼ 1, and the dotted curve was

calculated for another set of parameters, namely a ¼ 0.1,

b ¼ 0.71, and g ¼ 10.

Proton Conductivity of F0 7

Biophysical Journal 86(6) 1–16

MasterProof

DpH as the only driving force for protontransport: further evidence for the absenceof voltage-gating

Dimroth and co-workers have recently shown that the

operation in the synthesis direction of both the Na1-ATP

synthase of P. modestum and the H1-ATP synthase of E. colicannot be driven by an ion gradient alone but requires

a transmembrane voltage exceeding a certain threshold (Kaim

and Dimroth, 1998b, 1999; Dimroth et al., 2000). This

paralleled the previous observation of a voltage threshold of

the (oxidized) chloroplast enzyme (Junge, 1970; Junge et al.,

1970; Graber et al., 1977; Witt et al., 1977; Schlodder and

Witt, 1980), which could be replaced by chemical driving

force, DpH (Schlodder and Witt, 1981; Schlodder et al.,

1982). The latter now seems controversial. The data in Fig. 5,

B and D, clearly showed that proton-transfer through F0driven by a pH difference persisted, even after the electric

potential difference was dissipated in ,1 ms by the addition

of valinomycin. There was no voltage requirement and no

threshold for proton conduction by F0 itself. This was in line

with the above observation of the same electric relaxation

time observedwith 100%, 33%, and10%of the initial voltage.

It was obvious that F0 was not, by itself, voltage-gated.

The observed electrical gating in the intact enzymes from

E. coli and P. modestum was likely attributable to the

interaction of F1 with F0.

A kinetic model for proton conduction by F0

We investigated whether the magnitude, the Ohmic

behavior, and the weak pH-dependence of proton conduction

by F0 could be accommodated within the framework of the

presently assumed rotary mechanism of proton conduction

(Junge et al., 1997; Elston et al., 1998). The proton transfer

through the bare chloroplast F0 has been explained elsewhere

by a model where two proton-conducting half-channels were

connected by a (horizontally) rotating carrier (see Fig. 7 A for

an illustration). The fit of experimental data yielded a low

pK of 5–6 for the proton-accepting group of the input half-

channel and an alkaline pK of 8 for the output group

(Cherepanov et al., 1999). The weak pH-dependence be-

tween these two pK values follows from this model. The

proton-transfer rate is determined by the probability of pro-

ton binding to group A and proton release from group B.

When the ambient pH inside and outside are the same, as in

our experiments, the product probability is constant in this

interval. This feature explained the observed weak pH-

dependence of proton transfer through the bare F0. In ener-

gizedmembranes, on the other hand, the pKvalues of the relay

groups tend to match the ambient pH at the p- and n-sides ofthe couplingmembrane. This allows higher turnover numbers

of F0 even against backpressure (Cherepanov et al., 1999).

In an extension of the previous model (Cherepanov et al.,

1999), we assumed here that the pK values of the input and

output groups, located within the dielectric inside noncoaxial

proton-conducting half-channels as illustrated in Fig. 7 B, aresensitive to the membrane voltage in proportion to their

(dielectrically weighted) z-position in the membrane di-

electric. The respective potential profile of proton transfer is

schematically shown in Fig. 7 C.The main elements of the model were two noncoaxial

proton-conducting half-channels both considered to rapidly

FIGURE 6 Proton transfer through F0 as monitored by

two pH indicators reporting pH transients from either side

of the chromatophore membrane, (A, B) from the n-side

(Cresol red, 90 mM, at a wavelength of 575 nm), and (C,D) from the p-side (Neutral red, 26 mM plus 0.3% w/v

BSA, at 545 nm). Traces in parts A and C were recorded

in the absence, and traces B and D in the presence of

the ionophore valinomycin (2 mM). In the latter case the

transmembrane voltage was collapsed by the valinomycin-

mediated K1 current in,3 ms (see Fig. 2). The relaxation

of the pH difference was then entirely due to the entropic

driving force (DpH). The proton release from bc1 was

slower in the presence of valinomycin (compare to traces 2

in Figs. 6 C and 5 D). The medium contained 50 mM KCl,

5 mM MgCl2, 2 mM K4[Fe(CN)6], 2 mM KCN, 5 mM

dMF; pH was 7.9. Chromatophore stock was in 5 mM

MgCl2, 50 mM KCl, 10% sucrose. Traces 1 without, and

traces 2 with 3 mM oligomycin added; traces 2-1 ¼difference trace 6 oligomycin. Actinic flashes are marked

by arrows; black bars correspond to 0.002 pH units.

8 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

equilibrate with their adjacent bulk phase. In the simplest

case, each half-channel contained only one proton-binding

relay group (A at the acidic side of the membrane and B at the

basic side). In the absence of a transmembrane voltage, the

rate of proton binding to the groups A and B was assumed to

depend on the pH in the respective bulk phase in accordance

with the equations½AQ4�

Kon

A ¼ g � 1011 � ð10�pHA 1 10�pKA�14Þ;

Kon

B ¼ g � 1011 � ð10�pHB 1 10�pKB�14Þ; ðs�1Þ:

½AQ5� The first term in each equation denotes the proton flux

owing to the diffusion of H3O1 ions and the second term

describes the hydrolysis of neutral water (see Eigen, 1963).

The factor g was introduced to account for the accelerated

proton supply from surface buffering groups. (½AQ6� Addition of

penetrating pH buffers—glycylglycine and imidazole were

tested at concentration up to 0.2 M—caused no further

acceleration of proton transfer through F0; see Table 1. This

implies a rapid proton delivery along the inner chromato-

phore surface to F0 from many ionizable groups acting as

proton antennae. See e.g., Zhang and Unwin, 2002, for

a quantitative examination of surface proton conduction.)

The estimate of g was obtained by considering the strictly

Ohmic behavior of F0 with accuracy .3% (See Fig. 4).

Because the overall rate of proton transfer through F0 after

a saturating flash was � 3.5 3 103 s�1 (see above), the

linear relationship implied that at neutral pH the rate

constant konA (which was independent of DuuuH1 ) was .105

s�1, which corresponds to the g-factor .10 (see also the

discussion below). We used the latter value of g in the

modeling.

The rate constants of the reverse reactions (see the kinetic

scheme in Fig. 7 D) were calculated by the thermodynamic

equilibria of

Koff

A ¼ Kon

A � 10pHA�pKA ; Koff

B ¼ Kon

B � 10pHB�pKB ; ðs�1Þ:

The rate of proton exchange between F0 and the bulk

solutions was high and not limiting to the overall reaction

rate. The forward and reverse rate constants of the proton

transfer in the middle of membrane (m1 and m�) were

interconnected by the thermodynamic constraint of

m1 ¼ m� � 10pKA�pKB ðs�1Þ:Thus, there were only three independent kinetic parameters

in the model: the pK values of buffer groups in each of the

two half-channels, and the rate constant of the elementary

rotary step (the rotation of the cn-ring by the angle 2p/n),

which brings one proton into and another one off the ring in

contact with its respective access channel.

The potential energy of a transferred proton within F0 is

schematically plotted in Fig. 7 C. The transmembrane

electric field affects both the protonation state of the two

relay groups (A and B) in the two half-channels, and the rate

of proton transfer over the center of the membrane. The

magnitude of the first effect depends on the (dielectrically

weighted) relative depth of groups A and B in the membrane

(denoted a in Fig. 7 C, L is the full and 1 the relative

thickness of the membrane). The second effect depends on

the effective width of the barrier, denoted b in Fig. 7 C. Theabsolute height of the barrier is not important, if it is much

greater than the thermal energy kBT.A step of the transmembrane electric potential, magnitude

u (in Volts, positive at the A-side) shifts the pK values as

pK#A ¼ pKA 1auq=2:3 kBT;

pK#B ¼ pKB � auq=2:3 kBT;

where q is the charge of proton. The electric field also

changes the proton transfer rate across the middle of the

membrane, which is treated by the Nernst-Planck model

according to Hladky (1974) as

FIGURE 7 Functional model of F0. (A) Schematic illustration of F0operation. (B) Simple model for proton transfer; see text for details. (C)

Hypothetical energy profile for proton transfer; see text for details. (D)

Kinetic scheme.

Proton Conductivity of F0 9

Biophysical Journal 86(6) 1–16

MasterProof

m#1 ¼ m1 � e0:5�ð1�2aÞ�uq=kBT � f ðuÞ;m#� ¼ m� � e�0:5�ð1�2aÞ�uq=2kBT � f ðuÞ;

where the function F(u) depends on the shape of potential

barrier, which for a rectangular profile is

f ðuÞ ¼ ðbuqÞ=2 kBTÞ � sinhðbuq=2 kBTÞ�1

(see e.g., Bihler and Stark, 1997, for details). The kinetic

model shown in Fig. 7 B gives rise to a system of four

differential equations. Three equations describe the changes

of the population of the various states of the enzyme as

P00 ¼ �ðkonA 1 kon

B Þ � P00 1 koff

B � P10 1 koff

A � P01

P10 ¼ kon

B � P00 � ðkonA 1 koff

B 1m#�Þ � P10 1m#� � P01

1 koffA � P11:

P01 ¼ kon

A � P00 1m#� � P10 � ðkoffA 1 kon

B 1m#1 Þ � P01

1 koff

B � P11

These three differential equations have to be completed

by the normalizing condition

P00 1P10 1P01 1P11 ¼ 1:

Pij denotes the probability of protonation (index 1) and

deprotonation (index 0) of the relay groups in contact with

the left (left index) and the right (right index) access

channel.

The fourth differential equation describes the membrane

recharging

u ¼ FN0

Cm

½aðkonA ðP00 1P10Þ � koff

A ðP01 1P11Þ

1 konB ðP00 1P01Þ � koffB ðP10 1P11ÞÞ1 ð1� 2aÞðm1P01 � m�P10Þ�:

The solution of this system of four differential equations was

obtained by numerical integration.

Herein F denotes the Faraday constant, N0 the surface

density of F0 in the membrane (mol m�2), and Cm the electric

capacitance of membrane (F m�2). Because the average

diameter of chromatophores was�28 nm and they contained

a mean of �0.3 copies of F0 per vesicle, the mean surface

density of F0 in vesicles, which contained enzyme N0, was

�6.6 3 10�10 mol m�2. The membrane’s specific capaci-

tance was taken as 10�2 Fm�2. On modeling we took a value

of 70 mV for the average Dc jump in response to a single

saturating flash with the blocked bc1 complex. The fit

parameters pKA, pKB, and m1 were determined from the pH-

dependence of the F0 conductance by nonlinear minimiza-

tion. The final set of model parameters is summarized in

Table 2. The resulting theoretical pH-dependence of the

proton transfer rate in H2O is plotted in Fig. 3 by the solid

line. The obtained values pKA ¼ 6.1, pKB ¼ 10.0, and m1 ¼1 3 107 s�1 corresponded to the overall F0 turnover rate

m� ¼ m1 3 10pKA–pKB ¼ 1.26 3 103 s�1.

Fig. 5 A shows the normalized transients of the trans-

membrane voltage at pH 8.0 after a saturating (100%) and an

attenuated (33%) flash, the difference between them is

plotted in Fig. 5 B by the thin noisy line. This difference,

which is very small, characterizes the deviation from the

Ohmic behavior. The insert to Fig. 5 B shows the distribution

of the deviation between two traces, which did not reveal

a systematic bias from zero. The calculated mean root-square

dispersion of the difference was 3%.

The high signal/noise ratio of the traces allowed us to

impose constraints on the possible values of the parameters a,b, and g. Generally, the greater was a and the smaller b, the

faster was proton transfer at a given value of the trans-

membrane voltage, so the relaxation dynamics was faster

after saturating and slower after nonsaturating flashes

(positive nonlinearity). The effect of the parameter g was

opposite: because the diffusion of protons outside F0 was

voltage-independent, the proton diffusion at small values of g

slowed down the overall proton transfer (diffusional control

of the reaction) and caused thereby a negative nonlinearity in

the relaxation kinetics.

The dashed curve in Fig. 5 B shows the difference between

the transmembrane voltage decay after a saturating and an

attenuated flash calculated with the parameter g ¼ 1. This

curve revealed a substantial deviation from the Ohmic

behavior, which exceeded the experimental error. The

minimal value of g consistent with the experimental data

was�10, and this value was used in the following modeling.

The dotted curve in Fig. 5 B was calculated for a set of

parameters a ¼ 0.1, b ¼ 0.8, and g ¼ 10. It illustrates the

positive nonlinearity in the relaxation owing to the

accelerating effect by the transmembrane voltage. The

deviation again exceeded the experimental error, but had

the opposite sign.

We calculated the membrane discharge dynamics after

saturating and attenuated flashes using the kinetic model

described above and determined the range of possible values

of a and b consistent with an experimental error of ,3%.

Generally the relaxation rate was 10-fold more sensitive to

the variation of a (the buried position of groups A and B

inside the membrane) than to the variation of b (the form of

the potential barrier in the middle of the membrane). In the

two-dimensional phase space (a and b correspond to the

abscissa and ordinate axes of the insert to Fig. 5 A, respec-

TABLE 2 The kinetic and geometric fit parameters to the

data in Fig. 3 by the model illustrated in Fig. 7

pKA pKB m1 a b g

6.1 10.0 1.0 ½AQ8�107 s�1 #0.08 $0.6 10

10 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

tively) we determined the area where the maximal deviation

between the discharge curves after saturating and attenuated

flashes was being ,3% consistent with the experimental

error. The allowed space found by this method is shown in

the insert to Fig. 5 A by the uncolored upper-left area, and the

forbidden area inconsistent with the experimental data is

hatched. An example of the calculated difference between

two recharging traces with the maximal deviation of 3% is

shown in Fig. 5 B by the thick dashed line (the parameters

chosen were a ¼ 0.04, b ¼ 0.64, and g ¼ 10).

In practice, these findings implied that both proton-

conducting groups A and B were placed at the membrane/

water interface. The dielectrically weighted distance between

them had to be .0.86 (the maximal allowed value of a is

0.07). This maximal value of a corresponded, however, to an

infinitely sharp rectangular barrier in the middle of the

membrane (b¼ 1), physically unfeasible. A realistic value of

b is 0.6; it corresponds to an upper limit of a of �0.04. It is

worth noting, however, that the positive nonlinearity of the

electric field inside the membrane could be partially

compensated by the negative nonlinearity of the voltage-

independent diffusion outside the membrane, so we used

the value a ¼ 0.07 as an estimate of the buried position of

groups A and B inside the membrane. If the dielectric

permittivity was constant over the full width of the mem-

brane, say of 4 nm, the topological distance between A and B

would be .3.4 nm. The dielectric permittivity of the water-

filled cavities in membrane proteins may, however, strongly

differ from that in the membrane interior (see, e.g., a value of

30 in the ubiquinone binding pocket of bacterial reaction

center versus the value of 4 in the middle of membrane;

Cherepanov et al., 2000). In this case, the distance between

groups can be substantially smaller, on the order of 1.8 nm

(45% of the geometrical membrane thickness). It was ob-

vious though, that if either group was embedded more deeply

in the membrane dielectric, one had to expect a marked

deviation from an Ohmic behavior.

Our analysis reveals that the proton transfer to the

conserved carboxyl residue of subunit c, which is topolog-

ically located in the middle of the membrane, is mediated by

at least two residues located close to the membrane surface.

It remains open how this finding relates to the recent Cys-

mapping experiments of Angevine and Fillingame (2003),

which led these authors to postulate two water-accessible

half-channels in the F0F1-ATP synthase of E. coli. The relaygroups A and B may be found among those residues lining

these half-channels.

DISCUSSION

The Ohmic conductance of F0, itspH-dependence, kinetic H/D-isotope effect,and the absence of voltage-gating

In F0F1-ATP synthase the membrane-embedded F0 portion

conducts protons, and thereby it generates torque for the

synthesis of ATP by the F1 portion. We studied proton

conduction by F0 in chromatophores isolated from the purple

bacterium Rb. capsulatus. Rapidly rising voltage and/or pH

steps were generated by a flash of light. The subsequent

electric relaxation and the pH relaxation mediated by F0 were

detected by absorption transients of intrinsic (electrochro-

mic) pigments and of added pH-indicating dyes, respec-

tively. The vesicle size was so small that vesicles contained

less than a single conducting F0 molecule on the average,

namely 0.3. This was apparent from the fact that the elec-

trochemical relaxation was rapid only in a subset of vesicles

(26%); see Fig. 2. It was straightforwardly interpreted in

terms of a Poisson distribution of F0 over vesicles with

a mean of 0.3 and a frequency of 22% vesicles containing

one and 3% containing two copies of F0. This notion was

corroborated by the observation that the relative magnitude

of the rapid relaxation was larger in a preparation of chrom-

atophores with larger radius (see Feniouk et al., 2002, and

the text relating to Fig. 2 A of this article). When the small

contribution of vesicles containing more than one single

copy of F0 was neglected, we determined a unitary con-

ductance of 10.5 fS for the bacterial F0, which was equi-

valent to the translocation of 6500 H1/s at 100-mV driving

force.

In our related previous studies the average number of F0per vesicle was also small but larger than in this work

(Feniouk et al., 2002). Those particular preparations were,

however, prepared with a sonifier of smaller output power

(Branson W-15, Soest, The Netherlands). The average

diameter of the chromatophores used in the previous work

determined by TEM was 62.8 nm (data not shown), 1.6-fold

larger than that of the F1-depleted ones used here (see Fig.

1 A). The larger chromatophores contained approximately

one copy of F0F1 on the average, in good correspondence

with the F0 mean surface density calculated here.

We found that the electric relaxation through F0 was

almost mono-exponential and that the relaxation time was

practically independent of the magnitude of the initial

voltage step up to 70 mV. F0 behaved as an Ohmic conductor

with a linear current-voltage relationship. There was no

voltage-gating of F0. The conductance varied little (less than

threefold) as function of the pH over the wide range from 6 to

10. The specificity for protons over other cationic species

was very high (.107).

There was no kinetic H/D-isotope effect at pH above 8,

and an isotope effect of �2 was observed at pH 6 (Fig. 4 A).The appearance of the isotope effect could be attributed to

protonation of a group with the apparent pK of 7–8. A

possible candidate for this titratable group might be Glu61 of

the c-subunit. The pK of its counterpart in the E. coli enzyme

(cAsp61) has the apparent pK of 8 (Valiyaveetil et al., 2002).

An alternative explanation could be secondary isotope

effects: the general conformation of the enzyme might be

slightly altered by pH, and the H/D-isotope effect could

be inherent to the acidic conformation only. In this case

Proton Conductivity of F0 11

Biophysical Journal 86(6) 1–16

MasterProof

a smooth pH-dependence of the H/D isotope effect like that

in Fig. 4 A could be expected. The pH-dependence of both

the isotope effect and the Arrhenius activation energy (Fig.

4 B) points out that the rate-limiting step of proton trans-

port through F0 is different at pH 6 and 8. Further experi-

ments are necessary to clarify this point.

The above properties of the bacterial F0—the high

specificity, the weak pH-dependence, and the kinetic iso-

tope effect—were in a good agreement with those of the

chloroplast counterpart, where data were obtained over

a narrower pH interval, of 5.5 , pH , 8. (Althoff et al.,

1989). The very high proton specificity of F0 was attributable

to proton binding by acid/base group(s) as selectivity filter. It

excluded models with a contiguous water-wire without such

groups. The same property favored a Grotthuss-type

mechanism of proton transport, and it excluded the self-

diffusion of the proton in the form of H3O1 or OH�.

These properties of F0 were simulated based on the current

rotary mechanism with the multicopy c-ring carrying one

essential acid residue (Glu or Asp) in the middle of the

membrane and with one access channel at each side. We

assumed one proton relay group in each channel. The fit to

the data attributed pK values of�6 and�10, respectively, to

these relay groups because of the low dependence on the

pH. The Ohmic behavior called for a location of the relays

very close to their respective membrane/bulk interface. F0conducted protons irrespective of the nature of the driving

force, whether of electric (transmembrane voltage) or en-

tropic origin (pH difference): Both this property and the

Ohmic behavior under an electric driving force proved

the absence of voltage gating of F0. The DuuuH1 -gating of the

coupled enzyme in chloroplasts (Junge, 1970, 1987; Junge

et al., 1970; Graber et al., 1977; Witt et al., 1977; Schlodder

and Witt, 1980, 1981; Schlodder et al., 1982) and the

voltage-gating in E. coli and P. modestum (Kaim and

Dimroth, 1998a, 1999; Dimroth et al., 2000) is attributable to

the interaction of F1 with F0, but not to F0 proper.

The straightforward method used to obtain the unitary

conductance of one single copy of F0 in this work resolved

the discrepancy between previous reports from our lab. In

studies with spinach thylakoids we arrived at a similar

magnitude for the conductance (9 fS) under the then-

unproven assumption that any exposed F0 was conducting

(Schonknecht et al., 1986). Circumstantial evidence for only

a minority of conducting F0, which had led us to claim

a much higher conductance of up to 1 pS (Lill et al., 1986;

Althoff et al., 1989), can now be rejected. It also appears

conceivable that the similarly high conductance of 0.4 pS as

attributed to whole F0F1 in one study with bilayers formed by

the dip-stick technique (Wagner et al., 1989) might have

involved unspecific cation channels as observed with the

purified subunit c of F0 (Schonknecht et al., 1989).Extrapolating the data in this work on F0 to a driving

force of 200 mV (nearly sufficient for the maximum rate of

ATP synthesis by the coupled enzyme), one expects

a transport rate by F0 of 13,000 s�1. ATP synthesis by the

E. coli enzyme can reach a turnover number of 270 s�1

(Etzold et al., 1997), with coupled ATP hydrolysis in

Rhodospirillum rubrum at 320 s�1 (Slooten and Nuyten,

1984), and ATP synthesis in Rb. capsulatus at 250 s�1

(calculated from 900 mmol/mg BChl�h, Casadio et al., 1978,

if there was one ATPase per 1000 BChl; Packham et al.,

1978; Feniouk et al., 2002, and with a proton/ATP

stoichiometry of 3.3). Taking the proton/ATP stoichiometry

into account, which is probably 3.3 for yeast FoF1 (Stock

et al., 1999) and 4 (van Walraven et al., 1996; Turina et al.,

2003) or 4.7 (Seelert et al., 2000) for spinacia, these figuresimply the turnover of only �1000 H1/s by the coupled

enzyme, F0F1. This is one order-of-magnitude lower than

that of the exposed F0.

Comparison with the proton conductanceof gramicidin

Gramicidin is perhaps the best studied cation channel and

proton conductor. It is gated open when its two half-

spanning helices join tail-to-tail to fully span the membrane.

Cations are conducted along and with the internal water wire.

This is well established by studies on electro-osmosis and

streaming potentials. The proton, on the other hand, bypasses

the single-file transport (see Finkelstein and Andersen, 1981;

Levitt et al., 1978) and molecular dynamics simulation

(Skerra and Brickmann, 1987; Schumaker et al., 2000). The

proton conductance of the covalently linked gramicidin A

channels is proportional to the H1 concentration in the pH

range from 0 to 4 (Cukierman, 2000). Extrapolation to pH 8

yields a H1 conductance of 1 fS, 10-fold smaller than the H1

conductance of F0. In simulating the data on F0 we had to

assume that the supply to F0 was not rate-limiting, i.e., it was

by order-of-magnitude faster than expected by bulk diffusion

from a half-space to the channel mouth. It implied supply

from several buffering groups at the membrane surface

acting as proton donors. Recently, the lateral diffusion

coefficient along the lipid/water interface has been de-

termined to be 5.8 3 10�9 m2 s�1 (Serowy et al., 2003),

only twofold lower than in the bulk (9.3 3 10�9 m2 s�1),

but greater than for any other ion in the bulk phase and thus

compatible with a Grotthuss-mechanism involving surface

water (see Gutman and Nachliel, 1997). Proton supply in the

alkaline range is not critical, because water then acts as the

proton donor (see Kasianowicz et al., 1987; Gopta et al.,

1999). These features have been considered in the above

simulation.

CONCLUSIONS

The membrane portion, F0, of the F0F1-ATP synthase is

a ion-driven rotary engine. For the first time, its unitary

12 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

electric conductance was determined without ambiguity

concerning active and inactive copies of F0. We used

F1-depleted chromatophore vesicles from Rb. capsulatus thatwere so small as to contain only 0.3 copies of F0 on the

average. The conductance was constant as function of the

voltage, extremely proton-specific (.107), but only mildly

pH-dependent. The maximum conductance of 10.5 fS was

observed at pH 8 implying the translocation of 6500 H1 s�1

at 100-mV electric driving force and 13,000 at 200 mV.

Hence, F0, when operating without F1, turns over .10-fold

faster than it does in the coupled enzyme, F0F1. In other

words, F0 is not rate-limiting for the operation of F0F1, as

technically desirable. We found that F0, by itself, was not

voltage-gated. The observed electrical gating in the intact

enzymes from P. modestum and E. coli was likely attribut-

able to the interaction of F1 with F0. The nature of the gating

charge/dipole is still unknown.

In F0 the selectivity filter for protons over other cations

is highly specific, .107, both in the purple bacterium used

in this work and in thylakoids from green plants, as pre-

viously studied. The selectivity is still present but less

pronounced in organisms that operate on Na1 instead of H1.

We interpreted the properties of F0 in terms of the

current rotary model for proton conduction. This model is

based on two proton-conducting half-channels linking the

rotating ring of 10–14 copies of subunit c with the respec-

tive bulk phases. The observed Ohmic conduction implies

that the relay groups for protons that are involved in rate

limitation sample only a small fraction of the trans-

membrane voltage. We simulated such a behavior by two

relay groups located close to the respective membrane/

water interfaces. The slight variation of the conductance as

function of the pH implied a large difference between the

pKs of these groups (�6 and �10, respectively). This is

a testable prediction for structural studies aiming at high

resolution of the c-ring and its partners in the membrane,

the a- and b-subunits. That the proton conductance of F0was not limited by the supply of protons to the entrance

channel implied supply from buffers at the membrane

surface and from bulk water.

APPENDIX

The proton buffering capacity ofchromatophores

If the electrical and chemical components of DuuuH1 , Dc and DpH, were

discharged via F0 in isolation of each other, we observed different relaxation

times (see Fig. 5). This phenomenon has been previously observed in studies

on chloroplast F0 in spinach thylakoids (Schonknecht et al., 1986) and has

been interpreted as follows (Junge et al., 1992a,b). If Dc is the major driving

force of proton conduction, one observes the discharge of the electric capa-

citance (Cel), whereas the chemical capacitance (Cchem) of vesicles is

discharged if DpH is the major driving force. Presenting both in the same

physical dimensions, namely Farad � m�2½AQ7� , Cchem is related to the specific

proton buffering capacity of the vesicle, b ¼ �D½H1total�=DpH ðinmol �m�2Þ

as

Cchem ¼ F2

2:3RT3b;

wherein F denotes the Faraday constant, R the gas constant, T the

temperature, and b the specific buffering capacity, as defined above.

We described the proton flux through F0 by a two-step sequential kinetic

mechanism: A / B / C. The first step was the proton release by the bc1complex (in the presence of valinomycin, its relaxation time was 376 1 ms

as detected by Neutral red, see Fig. 6D). The second step (occurring with theunknown time t2) was the proton entry/escape through F0. The overall rate

of pH transients, as measured both by Cresol red outside and by Neutral red

inside chromatophores, was characterized by the summed time t11 t2 ¼ 52

6 5 ms. The proper time of DpH relaxation t2 was estimated thereby as 156

5 ms. This implied that the proton buffering capacitance of the vesicles at pH

8.0 was �3.5-fold greater than the electrical one. Taking the electric

capacitance of 1 mF cm�2, we calculated the specific proton buffering

capacitance of the internal surface of chromatophores b ¼ 2.1(60.7) 3

10�8 mol m�2. Because the surface density of RC in chromatophores is

�5.3 3 10�9 mol m�2 (Packham et al., 1978), this value corresponds to

eight buffer groups with pK ¼ 8 per RC. Assuming that all electron

equivalents, which were generated by the excited RCs after a saturating light

flash were completely utilized by the bc1 complexes, we calculated a pH

jump per saturating flash of 0.36 pH units. This estimate is in the range of

earlier ones, as obtained with 9-aminoacridine as a pH dye for

chromatophores of Rb. sphaeroides (0.32 pH unit; Saphon and Graber,

1978) and with Neutral red for chromatophores of Rb. capsulatus (0.86 0.2

pH unit; Mulkidjanian and Junge, 1994). The flash-induced pH jump in

bacterial chromatophores is by one order-of-magnitude greater than in

thylakoid membranes of green plants (0.06 units; Hong and Junge, 1983;

Junge et al., 1979). The difference could be attributed to the lower density of

proton pumps and larger amount of pH-buffering groups in thylakoids as

compared to the situation in bacterial chromatophores.

Valuable discussions with Profs. B. J. Jackson, B.-A. Melandri, and V. P.

Skulachev are appreciated.

This work has been supported in part by grants from the Deutsche

Forschungsgemeinschaft (Mu-1285/1, Ju-97/13, SFB 431-P15, 436-RUS-

113/210); by the Alexander von Humboldt Foundation; the Volkswagen

Foundation; the Leonhard-Euler Fellowship Program of Deutscher

Akademischer Austauschdienst; and the Fonds der Chemische Industrie.

REFERENCES

Altendorf, K., W. Stalz, J. Greie, and G. Deckers-Hebestreit. 2000.Structure and function of the F0 complex of the ATP synthase fromEscherichia coli. J. Exp. Biol. 203:19–28.

Althoff, G., H. Lill, and W. Junge. 1989. Proton channel of the chloroplastATP synthase, CF0: its time- averaged single-channel conductance asfunction of pH, temperature, isotopic and ionic medium composition.J. Membr. Biol. 108:263–271.

Angevine, C. M., and R. H. Fillingame. 2003. Aqueous access channels insubunit a of rotary ATP synthase. J. Biol. Chem. 278:6066–6074.

Auslander, W., and W. Junge. 1975. Neutral red, a rapid indicator for pHchanges in the inner phase of thylakoids. FEBS Lett. 59:310–315.

Baccarini-Melandri, A., H. Gest, and A. S. Pietro. 1970. A coupling factorin bacterial photophosphorylation. J. Biol. Chem. 245:1224–1226.

Bihler, H., and G. Stark. 1997. The inner membrane barrier of lipidmembranes experienced by the valinomycin/Rb1 complex: charge pulseexperiments at high membrane voltages. Biophys. J. 73:746–756.

Proton Conductivity of F0 13

Biophysical Journal 86(6) 1–16

MasterProof

Boyer, P. D. 1997. The ATP synthase—a splendid molecular machine.Annu. Rev. Biochem. 66:717–749.

Cao, N. J., W. S. Brusilow, J. J. Tomashek, and D. J. Woodbury. 2001.Characterization of reconstituted F0 from wild-type Escherichia coli andidentification of two other fluxes co-purifying with F0. Cell Biochem.Biophys. 34:305–320.

Casadio, R., A. Baccarini-Melandri, and B. A. Melandri. 1978. Limitedcooperativity in the coupling between electron flow and photosyntheticATP synthesis. FEBS Lett. 87:323–328.

Cherepanov, D. A., S. I. Bibikov, M. V. Bibikova, D. A. Bloch, L. A.Drachev, O. A. Gopta, D. Oesterhelt, A. Y. Semenov, and A. Y.Mulkidjanian. 2000. Reduction and protonation of the secondary quinoneacceptor of Rhodobacter sphaeroides photosynthetic reaction center:kinetic model based on a comparison of wild-type chromatophoreswith mutants carrying Arg/Ile substitution at sites 207 and 217 in theL-subunit. Biochim. Biophys. Acta. 1459:10–34.

Cherepanov, D. A., and W. Junge. 2001. Viscoelastic dynamics of actinfilaments coupled to rotary F-ATPase: curvature as an indicator of thetorque. Biophys. J. 81:1234–1244.

Cherepanov, D. A., A. Mulkidjanian, and W. Junge. 1999. Transientaccumulation of elastic energy in proton translocating ATP synthase.FEBS Lett. 449:1–6.

Clark, A. J., and J. B. Jackson. 1981. The measurement of membranepotential during photosynthesis and during respiration in intact cells ofRhodopseudomonas capsulata by both electrochromism and by permeantion redistribution. Biochem. J. 200:389–397.

Clayton, R. K. 1963. Absorption spectra of photosynthetic bacteria andtheir chlorophylls. In Bacterial Photosynthesis. H. Gest, A. San Pietro,and L. P. Vernon, editors. The Antioch Press, Yellow Springs, OH. 495–500.

Crofts, A. R., and C. A. Wraight. 1983. The electrochemical domain ofphotosynthesis. Biochim. Biophys. Acta. 726:149–185.

Cukierman, S. 2000. Proton mobilities in water and in differentstereoisomers of covalently linked gramicidin A channels. Biophys. J.78:1825–1834.

Dimroth, P. 2000. Operation of the F0 motor of the ATP synthase. Biochim.Biophys. Acta. 1458:374–386.

Dimroth, P., G. Kaim, and U. Matthey. 2000. Crucial role of the membranepotential for ATP synthesis by F1F0 ATP synthases. J. Exp. Biol.203:51–59.

Dimroth, P., H. Wang, M. Grabe, and G. Oster. 1999. Energy transductionin the sodium F-ATPase of Propionigenium modestum. Proc. Natl. Acad.Sci. USA. 96:4924–4928.

Eigen, M. 1963. Protonenubertragung, saure-base-katalyse and enzymati-sche hydrolyse. Teil I: elementarvorgange. Angew. Chem. 75:489–588.

Elston, T., H. Wang, and G. Oster. 1998. Energy transduction in ATPsynthase. Nature. 391:510–514.

Etzold, C., G. Deckers-Hebestreit, and K. Altendorf. 1997. Turnovernumber of Escherichia coli F0F1 ATP synthase for ATP synthesis inmembrane vesicles. Eur. J. Biochem. 243:336–343.

Feniouk, B. A., D. Cherepanov, W. Junge, and A. Mulkidjanian. 2001.Coupling of proton flow to ATP synthesis in Rhodobacter capsulatus:F1F0-ATP synthase is absent from about half of chromatophores.Biochim. Biophys. Acta. 1506:189–203.

Feniouk, B. A., D. Cherepanov, N. Voskoboinikova, A. Mulkidjanian, andW. Junge. 2002. Chromatophore vesicles of Rhodobacter capsulatuscontain on average one F0F1-ATP synthase each. Biophys. J. 82:1115–1122.

Fillingame, R. H., and O. Y. Dmitriev. 2002. Structural model of thetransmembrane F0 rotary sector of H1-transporting ATP synthasederived by solution NMR and intersubunit cross-linking in situ. Biochim.Biophys. Acta. 1565:232–245.

Fillingame, R. H., W. Jiang, O. Y. Dmitriev, and P. C. Jones. 2000.Structural interpretations of F0 rotary function in the Escherichia coliF1F0 ATP synthase. Biochim. Biophys. Acta. 1458:387–403.

Finkelstein, A., and O. S. Andersen. 1981. The gramicidin A channel:a review of its permeability characteristics with special reference to thesingle-file aspect of transport. J. Membr. Biol. 59:155–171.

Friedl, P., and H. U. Schairer. 1981. The isolated F0 of Escherichia coliATP-synthase is reconstitutively active in H1 conduction and ATP-dependent energy-transduction. FEBS Lett. 28:261–264.

Glasoe, P. K., and F. A. Long. 1960. Use of glass electrodes to measureacidities in deuterium oxide. J. Phys. Chem. 64:188–190.

Gopta, O. A., D. A. Cherepanov, W. Junge, and A. Y. Mulkidjanian. 1999.Proton transfer from the bulk to the bound ubiquinone QB of the reactioncenter in chromatophores of Rhodobacter sphaeroides: retardedconveyance by neutral water. Proc. Natl. Acad. Sci. USA. 96:13159–13164.

Graber, P., E. Schlodder, and H. T. Witt. 1977. Conformationalchange of the chloroplast ATPase induced by a transmembrane electricfield and its correlation to phosphorylation. Biochim. Biophys. Acta.461:426–440.

Gutman, M., and E. Nachliel. 1997. Time-resolved dynamics ofproton transfer in proteinaceous systems. Annu. Rev. Phys. Chem. 48:329–356.

Hille, B. 1984. Ionic Channels of Excitable Membranes. SinauerAssociated, Sunderland, Mass.

Hladky, S. B. 1974. The energy barriers to ion transport by nonactin acrossthin lipid membranes. Biochim. Biophys. Acta. 352:71–85.

Hong, Y. Q., and W. Junge. 1983. Localized or delocalized protons inphotophosphorylation? On the accessibility of the thylakoid lumen forions and buffers. Biochim. Biophys. Acta. 722:197–208.

Jackson, J. B. 1988. Bacterial photosynthesis. In Bacterial EnergyTransduction. C. Anthony, editor. Academic Press, London. 317–376.

Jackson, J. B., and A. R. Crofts. 1971. The kinetics of the light inducedcarotenoid changes in Rhodopseudomonas sphaeroides and their relationto electrical field generation across the chromatophore membrane. Eur. J.Biochem. 18:120–130.

Jiang, W., J. Hermolin, and R. H. Fillingame. 2001. The preferredstoichiometry of c-subunits in the rotary motor sector of Escherichia coliATP synthase is 10. Proc. Natl. Acad. Sci. USA. 98:4966–4971.

Junge, W. 1970. Critical electric potential difference for photophosphor-ylation. Its relation to the chemiosmotic hypothesis and to the triggeringrequirements of the ATPase system. Eur. J. Biochem. 14:582–592.

Junge, W. 1987. Complete tracking of transient proton flow through activechloroplast ATP synthase. Proc. Natl. Acad. Sci. USA. 84:7084–7088.

Junge, W. 1999. ATP synthase and other motor proteins. Proc. Natl. Acad.Sci. USA. 96:4735–4737.

Junge, W., G. Althoff, P. Jahns, S. Engelbrecht, H. Lill, and G.Schonknecht. 1992a. Proton pumps, proton flow and proton ATPsynthases in photosynthesis of green plants. In Electron and ProtonTransfer in Chemistry and Biology. A. Muller, H. Ratajczak, W. Junge,and E. Diemann, editors. Elsevier, Amsterdam, Netherlands. 253–72.

Junge, W., W. Auslander, A. J. McGeer, and T. Runge. 1979. The bufferingcapacity of the internal phase of thylakoids and the magnitude of thepH changes inside under flashing light. Biochim. Biophys. Acta. 546:121–141.

Junge, W., H. Lill, and S. Engelbrecht. 1997. ATP synthase: anelectrochemical transducer with rotatory mechanics. Trends Biochem.Sci. 22:420–423.

Junge, W., A. Polle, P. Jahns, G. Althoff, and G. Schonknecht. 1992b. Theelectrochemical relaxation at thylakoid membranes. In ElectrifiedInterfaces in Physics, Chemistry and Biology. R. Guidelli, editor.Kluwer Academic, Dordrecht, The Netherlands. 551–64.

Junge, W., B. Rumberg, and H. Schroeder. 1970. Necessity of an electricpotential difference and its use for photophosphorylation in short flashgroups. Eur. J. Biochem. 14:575–581.

Junge, W., and H. T. Witt. 1968. On the ion transport system ofphotosynthesis—investigation on a molecular level. Z. Naturforsch.23b:244–254.

14 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

Kaim, G., and P. Dimroth. 1998a. ATP synthesis by the F1F0 ATP synthaseof Escherichia coli is obligatorily dependent on the electric potential.FEBS Lett. 434:57–60.

Kaim, G., and P. Dimroth. 1998b. Voltage-generated torque drives themotor of the ATP synthase. EMBO J. 17:5887–5895.

Kaim, G., and P. Dimroth. 1999. ATP synthesis by F-type ATP synthaseis obligatorily dependent on the transmembrane voltage. EMBO J.18:4118–4127.

Kasianowicz, J., R. Benz, and S. McLaughlin. 1987. How do protonscross the membrane-solution interface? Kinetic studies on bilayermembranes exposed to the protonophore S-13 (5-chloro-3-tert-butyl-2#-chloro-4#-nitrosalicylanilide). J. Membr. Biol. 95:73–89.

Lascelles, J. 1959. Adaptation to form bacterichlorophyll in Rhodopseu-domonas sphaeroides: changes in activity of enzymes concerned inpyrrole synthesis. Biochem. J. 72:508–518.

Leslie, A. G., and J. E. Walker. 2000. Structural model of F1-ATPase andthe implications for rotary catalysis. Philos. Trans. R. Soc. Lond. B Biol.Sci. 355:465–471.

Levitt, D. G., S. R. Elias, and J. M. Hautman. 1978. Number of watermolecules coupled to the transport of sodium, potassium and hydrogenions via gramicidin, nonactin or valinomycin. Biochim. Biophys. Acta.512:436–451.

Lightowlers, R. N., S. M. Howitt, L. Hatch, F. Gibson, and G. Cox. 1988.The proton pore in the Escherichia coli F0F1 -ATPase: substi-tution of glutamate by glutamine at position 219 of the a-subunitprevents F0-mediated proton permeability. Biochim. Biophys. Acta. 933:241–248.

Lill, H., S. Engelbrecht, G. Schonknecht, and W. Junge. 1986. The protonchannel, CF0, in thylakoid membranes: only a low proportion of CF1-lacking CF0 is active with a high unit conductance (169 fS). Eur. J.Biochem. 160:627–634.

Linnett, P. E., and R. B. Beechey. 1979. Inhibitors of the ATP synthetasesystem. Meth. Enzymol. 55:472–518.

Melandri, B. A., A. Baccarini-Melandri, A. San Pietro, and H. Gest. 1970.Role of phosphorylation coupling factor in light-dependent protontranslocation by Rhodopseudomonas capsulata membrane preparations.Proc. Natl. Acad. Sci. USA. 67:477–484.

Melandri, B. A., A. Baccarini-Melandri, A. San Pietro, and H. Gest. 1971.Interchangeability of phosphorylation coupling factors in photosyntheticand respiratory energy conversion. Science. 174:514–516.

Mulkidjanian, A. Y., and W. Junge. 1994. Calibration and time resolutionof lumenal pH-transients in chromatophores of Rhodobacter capsulatusfollowing a single turnover flash of light: proton release by thebc1-complex is strongly electrogenic. FEBS Lett. 353:189–193.

Mulkidjanian, A. Y., M. D. Mamedov, and L. A. Drachev. 1991. Slowelectrogenic events in the cytochrome bc1-complex of Rhodobactersphaeroides: the electron transfer between cytochrome b hemes can benonelectrogenic. FEBS Lett. 284:227–231.

Muller, D. J., N. A. Dencher, T. Meier, P. Dimroth, K. Suda, H. Stahlberg,A. Engel, H. Seelert, and U. Matthey. 2001. ATP synthase: constrainedstoichiometry of the transmembrane rotor. FEBS Lett. 504:219–222.

Negrin, R. S., D. L. Foster, and R. H. Fillingame. 1980. Energy-transducingH1-ATPase of Escherichia coli. Reconstitution of proton transloca-ting activity of the intrinsic membrane sector. J. Biol. Chem. 255:5643–5648.

Noji, H., and M. Yoshida. 2001. The rotary machine in the cell, ATPsynthase. J. Biol. Chem. 276:1665–1668.

Packham, N. K., J. A. Berriman, and J. B. Jackson. 1978. The charg-ing capacitance of the chromatophore membrane. FEBS Lett. 89:205–210.

Panke, O., D. A. Cherepanov, K. Gumbiowski, S. Engelbrecht, and W.Junge. 2001. Viscoelastic dynamics of actin filaments coupled to rotaryF-ATPase: torque profile of the enzyme. Biophys. J. 81:1220–1233.

Panke, O., K. Gumbiowski, W. Junge, and S. Engelbrecht. 2000.F-ATPase: specific observation of the rotating c-subunit oligomerof EF0EF1. FEBS Lett. 472:34–38.

Sambongi, Y., Y. Iko, M. Tanabe, H. Omote, A. Iwamoto-Kihara, I. Ueda,T. Yanagida, Y. Wada, and M. Futai. 1999. Mechanical rotation of thec-subunit oligomer in ATP synthase (F0F1): direct observation. Science.286:1722–1724.

Saphon, S., and P. Graber. 1978. External proton uptake, internal protonrelease and internal pH changes in chromatophores from Rps.sphaeroides following single turnover flashes. Z. Naturforsch. 33C:715–722.

Saphon, S., J. B. Jackson, and H. T. Witt. 1975. Electrical potentialchanges, H1 translocation and phosphorylation induced by short flashexcitation in Rhodopseudomonas sphaeroides chromatophores. Biochim.Biophys. Acta. 408:67–82.

Schindler, H., and N. Nelson. 1982. Proteolipid of adenosinetriphosphatasefrom yeast mitochondria forms proton-selective channels in planar lipidbilayers. Biochemistry. 21:5787–5794.

Schlodder, E., M. Rogner, and H. T. Witt. 1982. ATP synthesis inchloroplasts induced by a transmembrane electric potential difference asa function of the proton concentration. FEBS Lett. 138:13–18.

Schlodder, E., and H. T. Witt. 1980. Electrochromic absorption changes ofa chloroplast suspension induced by an external electric field. FEBS Lett.112:105–113.

Schlodder, E., and H. T. Witt. 1981. Relation between the initial kineticsof ATP synthesis and of conformational changes in the chloroplastATPase studied by external field pulses.Biochim. Biophys. Acta. 635:571–584.

Schneider, E., and K. Altendorf. 1982. ATP Synthetase (F1F0) ofEscherichia coli K-12 high-yield preparation of functionalF0 by hydrophobic affinity chromatography. Eur. J. Biochem. 126:149–153.

Schneider, E., and K. Altendorf. 1987. Bacterial adenosine 5#-triphosphatesynthase (F1F0): purification and reconstitution of F0 complexes andbiochemical and functional characterization of their subunits. Microbiol.Rev. 51:477–497.

Schonknecht, G., G. Althoff, E. C. Apley, R. Wagner, and W. Junge. 1989.Cation channels by subunit III of the channel portion of the chloroplastH1 ATPase. FEBS Lett. 258:190–194.

Schonknecht, G., W. Junge, H. Lill, and S. Engelbrecht. 1986. Completetracking of proton flow in thylakoids—the unit conductance of CF0 isgreater than 10 fS. FEBS Lett. 203:289–294.

Schumaker, M. F., R. Pomes, and F. Roux. 2000. A combined moleculardynamics and diffusion model of single proton conduction throughgramicidin. Biophys. J. 79:2840–2857.

Seelert, H., A. Poetsch, N. A. Dencher, A. Engel, H. Stahlberg, and D. J.Mueller. 2000. Proton-powered turbine of a plant motor. Nature.405:418–419.

Senior, A. E., S. Nadanaciva, and J. Weber. 2002. The molecularmechanism of ATP synthesis by F1F0-ATP synthase. Biochim. Biophys.Acta. 1553:188–211.

Serowy, S., S. M. Saparov, Y. N. Antonenko, W. Kozlovsky, V. Hagen,and P. Pohl. 2003. Structural proton diffusion along lipid bilayers.Biophys. J. 84:1031–1037.

Skerra, A., and J. Brickmann. 1987. Simulation of voltage-driven hydratedcation transport through narrow transmembrane channels. Biophys. J. 51:977–983.

Slooten, L., and A. Nuyten. 1984. Steady-state kinetics of ADP-arsenateand ATP-synthesis in Rhodospirillum rubrum chromatophores. Biochim.Biophys. Acta. 766:88–97.

Sone, N., T. Hamamoto, and Y. Kagawa. 1981. pH dependence ofH1 conduction through the membrane moiety of the H1-ATPase (F0 3F1) and effects of tyrosyl residue modification. J. Biol. Chem. 256:2873–2877.

Stock, D., A. G. Leslie, and J. E. Walker. 1999. Molecular architecture ofthe rotary motor in ATP synthase. Science. 286:1700–1705.

Symons, M., C. Swysen, and C. Sybesma. 1977. The light-inducedcarotenoid absorbance changes in Rhodopseudomonas sphaeroides: an

Proton Conductivity of F0 15

Biophysical Journal 86(6) 1–16

MasterProof

analysis and interpretation of the band shifts. Biochim. Biophys. Acta.462:706–717.

Tsunoda, S. P., R. Aggeler, H. Noji, K. Kinosita, M. Yoshida, and R. A.Capaldi. 2000. Observations of rotation within the F0F1-ATP synthase:deciding between rotation of the F0 c-subunit ring and artifact. FEBSLett. 470:244–248.

Tsunoda, S. P., R. Aggeler, M. Yoshida, and R. A. Capaldi. 2001. Rotationof the c-subunit oligomer in fully functional F1F0 ATP synthase. Proc.Natl. Acad. Sci. USA. 98:898–902.

Turina, P., D. Samoray, and P. Graber. 2003. H1/ATP ratio of protontransport-coupled ATP synthesis and hydrolysis catalysed by CF0F1-liposomes. EMBO J. 22:418–426.

Valiyaveetil, F., J. Hermolin, and R. H. Fillingame. 2002. pH-dependentinactivation of solubilized F1F0 ATP synthase by dicyclohexylcarbodii-

mide: pKa of detergent unmasked aspartyl-61 in Escherichia coli subunitc. Biochim. Biophys. Acta. 1553:296–301.

van Walraven, H. S., H. Strotmann, O. Schwarz, and B. Rumberg. 1996.The H1/ATP coupling ratio of the ATP synthase from thiol-modulatedchloroplasts and two cyanobacterial strains is four. FEBS Lett. 379:309–313.

Wagner, R., E. C. Apley, and W. Hanke. 1989. Single channel H1 currentsthrough reconstituted chloroplast ATP synthase CF0-CF1. EMBO J. 8:2827–2834.

Witt, H. T., E. Schlodder, and P. Graber. 1977. Conformational change,ATP generation and turnover rate of the chloroplast ATPase analyzed byenergization with an external electric field. Dev. Bioenerg. Biomembr.1:447–457.

Zhang, J., and P. R. Unwin. 2002. Proton diffusion at phospholipidassemblies. J. Am. Chem. Soc. 124:2379–2383.

16 Feniouk et al.

Biophysical Journal 86(6) 1–16

MasterProof

QUERIES - biophysj36962q

[AQ1] Please note that throughout this article, ‘‘F0’’ was sometimes also rendered as ‘‘FO’’ and ‘‘Fo’’. These havebeen standardized throughout as ‘‘F0.’’ Please check this change. OK as done?

[AQ2] In this paragraph, are the two multidot characters indicating multipliers?

[AQ3] Please note that Biophysical Journal discourages the use of footnotes in text. Footnote 1 was placed here,between parentheses, next to its callout. OK as done?

[AQ4] Biophysical Journal requires multipliers to be set as multiplication symbols. For each of the six displayequations in which multidot(s) appear, please indicate if a multiplier is intended where the multidotcharacter is used. Thank you.

[AQ5] Please note that all equations had to be rekeyed, as they were inserted as graphics files and could not bestyled in that form. Please check all equations carefully to ensure their accuracy. Thank you.

[AQ6] Please note that Biophysical Journal discourages the use of footnotes in text. Footnote 2 was placed here,between parentheses, next to its callout. OK as done?

[AQ7] In this paragraph, are the two multidot characters indicating multipliers?

[AQ8] In Table 2, is the multidot character indicating a multiplier?

Biophysical Journal 2004This is your order form or pro forma invoice

(Please keep a copy of this document for your records)rev:11/01

Reprint Costs (Please see charge sheet)______ number of reprints ordered $__________Color in reprints, $262.00 per 100 copies $__________Each additional 100, add $82.00 $__________Each additional ship location, $32.00 $__________Free color publication does not apply toreprints.

Publication Fees (please see charge sheet for details)Page Charges:Members

$50 per page for pages 1-8 $__________$100 per page for pages 9-11 $__________$200 per page for pages 12 and above $__________

Nonmembers$70 per page for pages 1-8 $__________$100 per page for pages 9-11 $__________$200 per page for pages 12 and above $__________

Black and white reshoots, $9.00 each $__________Color reshoots, $60.00 each $__________Color Figures, $400.00 each $__________Number of approved free color figures _____

Total Amount Due $__________

Please note that authors of ‘‘New and Notable’’articles are exempt from publication fees.

The author will be notified of additional chargesdue to excessive corrections to the galley proofs.

Invoice Address (Invoice sent after publication)Name _______________________________________Institution _______________________________________Street _______________________________________

_______________________________________Department _______________________________________City _________________ State______ Zip________Country _______________________________________Phone ___________________ Fax ________________E-mail _______________________________________Purchase Order No. _________________________________

Shipping address (Cannot ship to PO Box)Name _______________________________________Institution _______________________________________Dept. _______________________________________Street _______________________________________

_______________________________________City _________________ State______ Zip________Country _______________________________________Quantity ___________________ Fax ________________Phone: Day ___________________ Evening ____________

Additional Shipping Address* (Cannot ship to PO Box)Name _______________________________________Street _______________________________________

_______________________________________City _________________ State______ Zip________Country _______________________________________Quantity ___________________ Fax ________________Phone: Day ___________________ Evening ____________*Add $32.00 for each additional shipping address

Credit Card Payment Details

Credit Card: ______ VISA ____ AMEX ___ MasterCardCard Number: __________________________________Expiration Date: __________________________________Signature: __________________________________Name as it appears on card: ________________________Billing address of cardholder: _________________________________________________________________________________________________________________________

Send your order form to:Dartmouth Journal ServicesAttn. Pat Bertozzi-Buck3 Archertown RoadBox 275Orford, NH 03777

Or fax: 603-353-9365

IT IS THE POLICY OF DJS TO ISSUE ONLY ONE INVOICE PER ORDER.

This form must be returned to Dartmouth Journal Services within 10 days of receipt,even if no reprints are ordered.

Author Name: ___________________________________________________________________________________Title of Article: ___________________________________________________________________________________Issue of Journal: ________________________________ Manuscript # ____________________________________No. of Pages: ________________________________ Color in Article: Yes / No (Please Circle)

Please include the journal name and manuscript number or reprint number on your purchase order or other correspondence

Enclosed: Institutional Purchase Order ___________________ Credit Card Payment Details _____________________Credit cards will be processed by DJS and a receipt will be sent with your invoice.

Signature: __________________________________________ Date: _______________________________Signature is required. By signing this form, the author agrees to accept the responsibility for the payment of reprints and/or allcharges described in this document.

Biophysical Journal2004 REPRINT AND PUBLICATION CHARGES

Black and White Reprint Prices

Minimum order is 100 copies. For articles longer than 32 pages,consult the Reprint Department at Dartmouth Journal Services.

Color in ReprintsAdd $262.00 per 100 copies (and $82.00 for each additional 100copies) to the cost of reprints if the article contains any color inaddition to black. For color orders greater than 500 copies, please callDartmouth Journal Services.

Publication FeesPlease note that ‘‘New and Notable’’ authors are exempt from publicationcharges.

CorrectionsPlease note that extensive substantive revisions will need tobe approved by the reviewing editor, resulting in the article appearingin a later issue or volume.

Authors will be granted up to an average of two corrections per page atno cost. Corrections, not including printer errors, made to the proofsthat exceed two per page will be charged to the author at a rate of$4.00 per correction. Figures that are replaced through no fault of thejournal office or the printer will also be charged to the author.

Page ChargesPage charges are $50 per page and $70 per page for members andnonmembers of the Biophysical Society respectively. Starting April 1,2003, page charges are tiered. Manuscripts whose corresponding

authors are members of the Biophysical Society at the time of originalsubmission pay a reduced $50 per page fee for pages 1–8, $100 perpage for pages 9–11, and $200 per page for pages 12 and over. Pagecharges for nonmember are $70 per page for pages 1–8, $100 per pagefor pages 9–11, and $200 per page for pages 12 and above. All authorsare expected to pay the page charges for articles they publish inBiophysical Journal. Failure to honor page charge obligations will affectfuture manuscript submissions. To apply for Society membership, visithttp://www.biophysics.org/members/memapp.pdf.

Figure ReshootsIf you are submitting figure reshoots, please add $9.00 for each blackand white reshoot and $60.00 for each color reshoot.

Articles Published with Color FiguresColor figures that, in the estimation of the Editorial Board with theadvice of the referees, are deemed essential to convey the science ofthe article will be printed free of charge to its authors, provided thecorresponding author of the paper is a current member of theBiophysical Society at the time of original submission. In general,color is not necessary for simple line drawings, x-y plots, bar graphs,or simple spectra. For papers submitted by nonmembers, the charge tothe authors for all color figures will be $400 per figure. A Societymember may request that color figures be included in one of his or herpapers in addition to those approved by the Editorial Board, but themember will be charged for all such color figures at the same rate asnonmembers.

ShippingUPS ground within the United States (1–5 days delivery) is included inthe reprint prices, except for orders in excess of 1,000 copies. Ordersshipped to authors outside the United States are mailed via anexpedited air service.

Multiple ShipmentsYou may request that your order be shipped to more than one location.Please be aware that it will cost $32.00 for each additional location.

DeliveryYour order will be shipped within 2 weeks of the journal print date.Allow extra time for delivery.

OrderingCredit card information or a signed institutional purchase order isrequired to process your order. Please return your order form, purchaseorder, and payment to:

Dartmouth Journal Services19 Archertown RoadBox 275Orford, NH 03777

Please direct all inquiries to Pat Bertozzi-Buck:E-mail: pbertozzi@dartmouthjournals.comPhone: (603) 353-9360 x105Fax: (603) 353-9365

International (includes Canada and Mexico)

> # ofpages

100 200 300 400 500

1-4 $275 $296 $323 $350 $380

5-8 $447 $500 $556 $609 $664

9-12 $612 $697 $782 $865 $950

13-16 $777 $889 $1,003 $1,116 $1,232

17-20 $947 $1,087 $1,233 $1,373 $1,514

21-24 $1,116 $1,286 $1,459 $1,627 $1,800

25-28 $1,287 $1,483 $1,688 $1,885 $2,083

29-32 $1,458 $1,680 $1,919 $2,139 $2,367

This order form and prepayment or signedinstitutional purchase order must be

returned to Dartmouth Journal Services within10 days of receipt.

Domestic (USA Only)

> # ofpages

100 200 300 400 500

1-4 $254 $273 $294 $314 $334

5-8 $423 $459 $499 $535 $576

9-12 $580 $637 $696 $756 $814

13-16 $733 $813 $892 $927 $1,050

17-20 $890 $988 $1,090 $1,189 $1,289

21-24 $1,047 $1,165 $1,290 $1,407 $1,525

25-28 $1,205 $1,343 $1,488 $1,624 $1,762

29-32 $1,363 $1,518 $1,688 $1,843 $1,999

MasterProof

QUERIES - biophysj36962q

[AQ1] Please note that throughout this article, ‘‘F0’’ was sometimes also rendered as ‘‘FO’’ and ‘‘Fo’’. These havebeen standardized throughout as ‘‘F0.’’ Please check this change. OK as done?

[AQ2] In this paragraph, are the two multidot characters indicating multipliers?

[AQ3] Please note that Biophysical Journal discourages the use of footnotes in text. Footnote 1 was placed here,between parentheses, next to its callout. OK as done?

[AQ4] Biophysical Journal requires multipliers to be set as multiplication symbols. For each of the six displayequations in which multidot(s) appear, please indicate if a multiplier is intended where the multidotcharacter is used. Thank you.

[AQ5] Please note that all equations had to be rekeyed, as they were inserted as graphics files and could not bestyled in that form. Please check all equations carefully to ensure their accuracy. Thank you.

[AQ6] Please note that Biophysical Journal discourages the use of footnotes in text. Footnote 2 was placed here,between parentheses, next to its callout. OK as done?

[AQ7] In this paragraph, are the two multidot characters indicating multipliers?

[AQ8] In Table 2, is the multidot character indicating a multiplier?